Infectious pneumonia is among the most common infections in human. An appropriate in vivo model is critical for understanding disease pathogenesis and testing the efficacy of novel therapeutics. With this murine oropharyngeal aspiration pneumonia model, one can examine the pathogenesis and new treatments against these deadly infections.
Murine infection models are critical for understanding disease pathogenesis and testing the efficacy of novel therapeutics designed to combat causative pathogens. Infectious pneumonia is among the most common infections presented by patients in the clinic and thus warrants an appropriate in vivo model. Typical pneumonia models use intranasal inoculation, which deposits excessive organisms outside the lung, causing off-target complications and symptoms, such as sinusitis, gastritis, enteritis, physical trauma, or microparticle misting to mimic aerosol spread more typical of viral, tuberculous, or fungal pneumonia. These models do not accurately reflect the pathogenesis of typical community- or healthcare-acquired bacterial pneumonia. In contrast, this murine model of oropharyngeal aspiration pneumonia mimics the droplet route in healthcare-acquired pneumonia. Inoculating 50 µL of the bacteria suspension into the oropharynx of anesthetized mice causes reflexive aspiration, which results in pneumonia. With this model, one can examine the pathogenesis of pneumonia-causing pathogens and new treatments to combat these diseases.
Lower respiratory infection is the world's deadliest communicable disease and the most common cause of death in developing countries1. Globally, these infections account for more than 3.2 million deaths1. In addition, nosocomial pneumonia is among the most common and deadly forms of healthcare acquired infections, and is caused by the most antibiotic-resistant pathogens2,3. The typical route of acquisition of bacterial pneumonia for both community-acquired and nosocomial pneumonia is the aspiration of oropharyngeal contents into the alveoli. Murine models used to study these diseases often use intranasal inoculation4, depositing much of the bacteria outside the lung, causing off-target complications and symptoms like sinusitis and physical trauma, which are incongruent with the disease progression in human that the models were designed to emulate. Other models may use inhalation chambers and micromisting devices, which more accurately mimic viral, tuberculous, and fungal pneumonias, but do not accurately recapitulate the normal route of acquisition for typical bacterial pneumonias.
The murine oropharyngeal aspiration pneumonia model may be utilized to simulate the natural route and pathogenesis of bacterial pneumonia. By inoculating 50 µL of the bacterial suspension into the oropharynx of anesthetized mice using a pipette, reflexive aspiration ensues, which results in infectious pneumonia. Using this model, one can examine the pathogenesis of pneumonia-causing pathogens and new treatments to combat these diseases with a higher fidelity model, more analogous to aspiration pneumonia infections observed in human. Additionally, unlike similar models that infect through the oral cavity5,6, this model ensures that the full inoculum reaches the lungs instead of the gut, where it can cause off-site inflammation and infections, such as gastritis and enteritis. Finally, unlike another published model that requires a laryngoscope and inoculates through the trachea7, this model does not obstruct the airway with a gavage needle and does not require injection for inoculum delivery. Instead, inoculation relies on the natural aspiration reflex of the mouse.
All procedures involving animals must be approved by the researcher’s Institutional Animal Care and Use Committee (IACUC).
1. Preparation of Bacterial Inoculum
2. Anesthetizing Mice
3. Inoculating/Infecting Mice
4. Monitoring Disease Progression
Note: Due to animal suffering, various indicators should be used to indicate when mice become moribund; euthanasia should be performed following this determination, in accordance with a previously approved IACUC protocol; various markers of moribundity include body temperature, weight loss, appearance, gait, and other biomarkers that can be obtained from blood (e.g., via iSTAT)9,10,11,12,13,14,15,16.
By carefully following the protocol, reproducible and robust data can be easily obtained. It is critical to strictly adhere to one's customized inoculum preparation protocol for experiments to be compared to one each other. It is also important to properly handle mice during the infection procedure. Be sure to place mice into an anesthesia chamber devoid of isoflurane. Mice will panic if they are placed into a chamber that has been pre-filled with isoflurane and may experience excess stress, which can possibly compromise experimental results. After securing the container lid, slowly introduce isoflurane by incrementally increasing its concentration from 0% to 4% (v/v); hastily administering isoflurane can also cause mice to panic. Once the mice become unconscious, reduce the isoflurane concentration to 2 – 3% (v/v) and allow them to remain in the chamber for an additional few minutes. At this point, mice are ready to be infected (Figure 1).
Remove a mouse from the anesthesia chamber and suspend it by its top incisors to allow access to the tongue (Figure 2). Use blunt-ended forceps to gently pull out the tongue (Figure 3) then transfer the forceps-held tongue to a sterile, gloved fingers (Figure 4) to prevent trauma to the mouse's tongue. With the tongue still outside the mouth, transfer the 50-µL inoculum to the oropharynx (Figure 5). Here, it is very important to only deliver liquid by pipetting to the first stop on the micropipette. Continuing to the second stop can introduce a large bubble into the inoculum that interferes with the infection.
Once the infection is complete, there are a myriad of options for testing mice. For pathogenesis studies, one can compare uninfected to infected tissue (Figure 6). If analyzing the effectiveness of novel therapeutics, one can remove the lungs and have them sectioned and stained. This can be done at one time point or at multiple time points to show disease progression (Figure 7).
Another option is to assess the bacterial burden in the lungs of infected mice. This is done by removing the lungs, transferring to a vial containing a known volume of sterile PBS, then homogenizing with a tissue homogenizer (rinsing between each sample to prevent cross contamination). Plating serial dilutions of the lung homogenate on nutrient-rich agar allows for calculating the CFUs/mL for each lung homogenate and subsequently CFUs/mg lung tissue (Figure 8). This, too can be done at one time point or at multiple time points to show disease progression.
There are countless other assays that can be performed, including cytokine analyses (Figure 9), sepsis biomarkers by iSTAT (Figure 10), flow cytometry to investigate cell surface markers, cellular profiling, RNAseq, etc.
Figure 1: Determining LD100. It is necessary to infect mice with various inocula concentrations to determine the LD100. Here, the inoculum 2 × 108 CFUs/mouse is too high, 5 × 107 and 2 × 107 are too low, and 1 × 108 is just right.
Figure 2: Hang mice by top incisors. After the mice are anesthetized, remove one mouse from the induction chamber, (A) use forceps to pull out a loop of string secured 20 – 30 cm above the work surface, (B) move the string behind the mouse's top incisors, (C) ensure the string is secured behind top incisors, and (D) let the mouse hang by its top incisors on the loop of string.
Figure 3: Pull tongue out of mouth with forceps and transfer grip to gloved fingers. After hanging the mouse by its top incisors, (A) use forceps to gently grasp the mouse's tongue, (B) gently pull out the tongue, (C) carefully transfer the tongue from forceps to gloved fingers to prevent trauma to the mouse's tongue, and (D) hold the tongue in place with gloved fingers. Be sure to apply enough pressure to prevent the tongue from slipping back into the mouth but not so much pressure that trauma ensues. The tongue should be held outside of the moth and to the side to allow for micropipette access in the next step. At this point, the mouse is ready for infection.
Figure 4: Infect the mouse. Maintaining the tongue outside the mouth, use a micropipette to transfer the 50-µL inoculum to the oropharynx. Place the pipet tip inside the mouth at the back of the tongue and dispense the bacterial suspension, only going to the first top on the micropipette; do not go to the second stop as this can introduce a large bubble into the inoculum that can interfere with complete aspiration.
Figure 5: Micrograph of healthy (uninfected) vs infected lung tissue. Hematoxylin and eosin (H&E) staining of lung sections before and after oropharyngeal aspiration of A. baumannii, which recapitulates the route of ventilator-associated pneumonia (VAP), resulting in death typically within 1 – 3 days and substantial alveolar inflammation as seen here.
Figure 6: Assess bacterial burden; reprinted and modified with permission14. After infecting mice, remove lungs by dissection following successful euthanasia in compliance with the applicable IACUC protocol. Collect the mass of the lung tissue, transfer to a vial containing a known volume (2 – 5 mL) of sterile PBS, then homogenize with a tissue homogenizer. Be sure to rinse the homogenizer with ethanol and sterile PBS in between each sample to prevent cross contamination. Perform serial dilutions of the lung homogenate and plate various dilutions on nutrient-rich agar and incubate appropriately for the chosen pathogen. Calculate the CFUs/mL for each lung homogenate based on CFUs/plate × dilution, and then divide by the mg lung tissue/mL lung homogenate. This can be done at one time point or at multiple time points to show disease progression. Medians are displayed with error bars representing interquartile ranges. *p <0.05 Treated vs. Untreated group.
Figure 7: Micrograph of infected lung tissue, untreated vs treated; reprinted and modified with permission14. After infecting mice, remove lungs by dissection following successful euthanasia in compliance with the applicable IACUC protocol. Appropriately preserve the tissue (e.g., immersed in Optimal Cutting Temperature gel and frozen at -80 °C) then send to pathology lab or other proficient individual for sectioning and staining. This can be done at one time point or at multiple time points to show disease progression.
Figure 8: Cytokine analysis; reprinted and modified with permission14. To analyze circulating cytokines, procure sufficient blood (e.g., 50 µL via tail nicking), allow to coagulate for 30 min at room temperature, centrifuge at 1,000 × g for 10 min at 4 °C, and collect serum supernatant. The serum can then be analyzed by Luminex multiplex for cytokines. This can be done at one time point or at multiple time points to show disease progression. Medians are displayed with error bars representing interquartile ranges. *p <0.05 treated vs. untreated group at same time-point.
Figure 9: Sepsis biomarkers; reprinted and modified with permission14. To analyze sepsis biomarkers, procure 75 µL blood (e.g., via tail nicking) and quickly transfer to an iSTAT cartridge that tests for desired analytes. This can be done at one time point or at multiple time points to show disease progression. Medians are displayed with error bars representing interquartile ranges. *p <0.05 treated vs. untreated group at same time-point.
To be sure, mice are not miniature humans. Results obtained from mouse models must be considered in context and subsequently interpreted for applicability to humans, based on differences and similarities between the two species6. It is also important to choose the appropriate mouse strain as certain are more susceptible to some infections than others; the same applies to the pathogen strain of choice16.
It is essential to perform infections in an exacting and highly reproducible way. Inocula can be lethal to mice at one value yet harmless at even 90% of that value. Thus, it is imperative that all conditions listed in this protocol are reproduced in the exact same way, particularly when preparing the inoculum and when infecting. Additionally, each pathogen will cause disease at a unique inoculum. Therefore, it is necessary to perform pilot experiments to determine the appropriate inoculum for each strain of pathogen and in each strain of mouse.
With Gram-negative bacteria, an inoculum of ≤5 × 108 CFUs/mouse is sufficient to cause death in virulent strains. It is advised to not use strains that are non-lethal at ≤1 × 109 CFUs/mouse because they suggest dubious relevance to pathogenesis based on sheer quantity of material being placed into the lungs16.
Compared to others, there are very few technical complications for the oropharyngeal aspiration pneumonia model and reproducibility is far greater. Only one minor complication of the model results when the surface tension around the 50-µL inoculum placed in the oropharynx allows for nasal respiration, precluding aspiration-the cause of infection. In this case, simply pinching the nares with forceps compels reflexive aspiration through the mouth and subsequent inhalation of the inoculum to induce infectious pneumonia. This is, however, a technical error by the technician, which is easily avoided with minimal practice.
In terms of its relevance to human disease, the one limitation of this model, like other models of Gram-negative bacterial infections, is the rapidity of the progression of the disease. Humans are rarely infected with a large bolus of a bacterial suspension, which is why mice progress to disease so rapidly. Nevertheless, all FDA-approved treatments undergo such translational research screenings in animal models to assess their therapeutic potential before moving on to clinical trials in humans. Ironically, the rapidity of the model is also an attractive feature, since it can provide clarity on therapeutic efficacy within a brief period of time.
No murine model is perfect, but this is a model with the ability to faithfully emulate the route of infection and pathogenesis of human diseases and assess the effectiveness of potential therapeutics, thereby allowing for the rapid translation of novel therapies that are desperately needed in the clinic.
The authors have nothing to disclose.
This work was supported by the National Institute of Allergy and Infectious Diseases at the National Institutes of Health [Grant Numbers R01 AI117211, R01 AI130060, R21 AI127954, and R42 AI106375 to BS] and US Food and Drug Administration [Contract HHSF223201710199C to BML].
Agar | BD | 214530 | Combine with TSB to make TSA |
Beads, Borosilicate Glass | Kimble | 135003 | Sterilize by baking or autoclaving before each use |
Beaker, 250 mL | Pyrex | 1003 | Used during precise aliquoting of concentrated bacterial inocula |
Centrifuge | Sorvall | ST 40R | Capable of 4,000×g at 4°C |
Chamber for Anesthesia | Kent Scientific Corporation | VetFlo-0720 | Accommodates up to 5 mice |
Cryomold, Intermediate Size | Sakura Tissue-Tek | 4566 | Disposable vinyl specimen molds, 15×15×5 mm |
Dental Floss | Oral-B | 37000469537 | Tie to stable post approx. 6" above table height |
Forceps | VWR | 82027-440 | Used to gently pull tongue out of mouse's mouth |
Homogenizer for Lung Tissue | Omni International | TM125-115 | Autoclave before first use; rinse between samples |
Isoflurane for Anesthesia | Abbott | 10015516 | Alternative drug can be used; modify procedure accordingly |
iSTAT Cartridge | Abbott | 03P79-25 | Various cartridges are available to suit your needs |
Ketamine, 100 mg/mL | Western Medical Supply | 4165 | Dilute 1:10 in PBS to 1 mg/mL and combine with Xylazine at 1 mg/mL |
Ointment for Eyes | Akorn | Tears Renewed | Avoid touching eye with tip of dispenser |
Optimal Cutting Temperature (O.C.T.) Compound | Fisher Scientific | 23-730-571 | Used to freeze lung samples at -80 °C to prepare for pathology sectioning |
Petri Dish | VWR | 25384-302 | Polystyrene, disposable, sterilized, 100×15 mm |
Phosphate-Buffered Saline (PBS) | Corning | 21-031-CM | Dulbecco's PBS without calcium and magnesium |
Pipette Tips, 200-μL | VWR | 10017-044 | Autoclave before use |
Pipetter, 200-μL | Gilson | Pipetman P200 | Autoclave and calibrate before use |
Spreader, Bacterial Cell | Bel-Art | F377360006 | Sterilize by baking or autoclaving before each use |
Stir Bar, Magnetic, 7.9 mm Diameter × 38.1 mm Length | VWR | 58948-150 | Used for stiring concentrated bacterial inocula during aliquoting |
Stir Plate, Magnetic | Corning | PC-620D | Used for stiring concentrated bacterial inocula during aliquoting |
Tryptic Soy Broth (TSB) | BD | 211822 | Combine with Agar to make TSA |
Vial, Conical, Sterile, 50 mL | Corning | 431720 | Used for preparing bacterial inocula |
Vial, Conical, Sterile, 500 mL | Corning | 431123 | Used to concentrate inocula for preparing frozen inocula |
Vial, Cryogenic, 2.0 mL | Corning | 430659 | Used for cryogenic storage of concentrated bacterial inocula |
Xylazine, 20 mg/mL | Akorn | AnaSed Injection | Dilute 1:20 in PBS to 1 mg/mL and combine with Ketamine at 1 mg/mL |